It is much easier to avoid bubbles if you fill the space with enough solution that it spills over the top as you put the comb in.

Attach binder clips to help hold the comb in while drying. One in either side of the casting stand clamp.

Leave for 1 hour while polymerization occurs.

Can store for a few weeks in the fridge. Leave comb in, and wrap in a wet paper towel and cling wrap.

binder clips squeeze the glass to the comb. Put them as far down as they go.

Consolidated/Pictorial Protocol

SDS casting protocol

Sample Prep

use 5-8 or even up to 20 ug protein per well. (for Mini Protean with 10 well comb)

If biomass is not limiting, prep a significant excess of protein, which will allow you to re-run if your gel turns out poorly for any reason.

If protein normalizing:

lyse cultures by sonication. Use 20 1 second pulses while tube is in ice, let the samples rest 5 minutes on ice, then sonicate again. Wipe sonicator stick between uses. Don't touch the tip while it is on.

sonication allows you access to un-dyed lysed sample, unlike boiling with sample loading buffer as is described below.

Mistakes to Be Careful About

using an "old" APS solution when making gels. The 10% weight/volume APS should be made fresh each time for best results. Don't use a solution that is more than a month or so old. The gels won't polymerize.

not mixing the liquid gel mixture enough

letting the gel dry too long after pouring the stacking gel (comb step)

the very edges can shrivel up, which becomes a problem when you try to use those edge lanes

taking the gel off the casting stand before it has polymerized

entire sample will leak out

sample sloshing out of the well you are using into a neighboring well

using too much beta-mercaptoethanol in your sammple buffer

should have < 1% beta-mercaptoethanol in the mix after you add sample buffer to the

too much reduction of cysteines is bad: will alter structure and even cleave proteins.

not having the lid to the running unit on all the way. --> poor electrical contact & blurry bands

note: Coomassie Brilliant Blue G-250 differs from Coomassie Brilliant Blue R-250 by the addition of two methyl groups. We use the G form. Read more about the R form here or at the bottom of this page.

Destaining Buffer:

30% methanol, 10% acetic acid, water

some labs use much less methanol & acetic acid; some use plain water.

Janet rinses in plain water before using our destianing buffer.

Acrylamide toxicity

Acrylamide is toxic to your nervous system, and may be a carcinogen. The unpolymerized form is toxic, but the polymerized form is much less toxic. ALWAYS wear gloves and wipe up spills - once the solution drys, the dust can be inhaled. Interestingly, fried starchy/sugary foods naturally contain acrylamide, too.

more than you want to know about acrylamide toxicity can be found here

Ladder

PageRuler protein gel legend

Other ladders:

BioRad protein standards

Combs/Loading Volumes

We currently only have the mini-protean gel running boxes. -JM 10/2012

SDS combs and loading volumes

Questions/Answers/Facts

What do all of the reagents do?

SDS

In loading buffer and often in gels (not necessary to include in gels; can be used in sample buffer)

APS & TEMED

polymerize acrylamide

glycine

carry charge in the opposite direction as the negatively-charged SDS-covered proteins

methanol

in wash buffer

glycerol in loading buffer: helps sample sink

Bromphenol Blue

Is it important to degas my water + buffer + acrylamide mix before adding the APS and TEMED, as the manuals recommend?

"We recommend that you degas the solutions to get rid of nitrogen. Sometimes, the nitrogen may come out of solution and form bubbles in your gels. " -BioRad customer support 10/26/2012 JM

"Proper degassing and filtering of the casting solution is critical for both reproducibility of the polymerization (oxygen removal)" (link)

This manual suggests the consequence is poor polymerization. If you don't experience poor polymerization, maybe you don't need to worry about degassing...?

Also note this is not the justification customer support provided. (see above)

Why shouldn't we overlay the gels with butanol or isoporpanol as they polymerize?

"We do not recommend using butanol/isopropanol because these may degrade the glue on the spacer plates and the plastic of the casting frame." -BioRad customer support 10/26/2012 JM

Should I soak my gel or run them empty before use?

They say no: "I have not heard that soaking the gels in water will improve the run. Talking with my colleagues, we thought it would have been detrimental because the tris and chloride ions will diffuse out during this time and in theory, should have made your gel run poorly."

JM had a gel that was only loaded with two lanes of ladder run funny. The ladder bands ran fine until 1/2 way through the gel, after which they halted and stacked on top of one another. Immediately after, I could see a 1 cm band in the gel that refracted light differently. This band disappeared after a while of soaking, leading me to believe it was a buildup of one of the buffer compounds. I soaked this gel overnight in water, loaded ladder in an unused adjacent well, and it ran perfectly without bunching up. The ladder ran fine until it reached ~ 1/2 way down, where it bunched up. A ~ 1 cm horizontal strip of the gel refracted light differently, but this band went away over time. It was repeatable with gels in the same batch, and with a batch Amanda prepped (though hers were less dramatic).

We currently believe it is best to soak gels overnight or run them in buffer for ~ 1 hour at 200V before use. BioRad tech support suggested that if we chose to soak (which they don't officially recommend), we should do so with You can use 0.375 M Tris, pH 8.8. This is the concentration of the storage buffer used in BioRad's precast gels. -JM 10/29/2012

Potential problems with soaking or pre-running gels:

You might ruin the stacking nature of the gel by alterning the buffer within the acrylamide matrix. This essentially converts your gel to a continuous buffer gel system, known to give more blurry bands.

Is it ok to re-use my electrode buffer?

Many people re-use it up to ~ 20 times, however, you should know that the manuals recommend single use and understand why. This may help.

How long can I store my acrylamide gels?

"Tris-HCl resolving gels are prepared at pH 8.6–8.8. At this basic pH, polyacrylamide slowly hydrolyzes to polyacrylic acid, which can compromise separation. For this reason, Tris-HCl gels have a relatively short shelf life. In addition, the gel pH can rise to pH 9.5 during a run, causing proteins to undergo deamination and alkylation. This may diminish resolution and complicate postelectrophoresis analysis." (reference)

note: commerical Tris gels have a shelf life of 6 months to 1 year. Amanda & Ceci don't keep hand-poured gels for more than a few weeks. (JM 11/2012)

It is possible the commercial gels have other means of extending shelf life.

The G-250 form the colloidal particles in an aqueous solution. This is an advantage for staining a gel because the colloids tend not to stain the gel matrices, reducing the background problem. When the colloids come close to the proteins, the dye molecule is removed from the colloids by the nearby proteins due to the higher affinity of proteins to the dye.

R-250, on the other hand, doesn't form the colloids. Rather, an individual dye molecule is dispersed in a solution. Therefore, the dye molecules can interact not only with proteins but with gel matrices freely, creating the background staining issue.

Other Tips

Excess salt in SDS-PAGE samples causes fuzzy bands and narrowing of gel lanes toward the bottom of the gel

If the ionic strength is very high, no bands will appear in the lower part of the gel (a vertical streak will appear instead) and the dye front will be wavy instead of straight. Deionize any sample with a total ionic strength over 50 mM using columns such as Micro Bio-Spin™ columns, which contain 10 mM Tris at a pH suitable for SDS-PAGE. (source)

Success or failure of any protein analysis depends on sample purity.

Interfering substances that can negatively impact SDS-PAGE include salts, detergents, denaturants, or organic solvents (Evans et al. 2009). Highly viscous samples indicate high DNA and/or carbohydrate content, which may also interfere with PAGE separations. In addition, solutions at extreme pH values (for example, fractions from ion exchange chromatography) diminish the separation power of most electrophoresis techniques. Use one of the following methods as needed to remove these contaminants:

Protein precipitation — the most versatile method to selectively separate proteins from other contaminants consists of protein precipitation by trichloroacetic acid (TCA)/acetone followed by resolubilization in electrophoresis sample buffer. A variety of commercial kits can simplify and standardize laboratory procedures for protein isolation from biological samples

Polyacrylamide is oxidative, and so disulfide bonds may be re-formed after proteins enter the gel

also, oxidation of agents used to reduce any disulfide bonds in the original sample may introduce artifactual new components in the gel pattern. (source)

SDS can form micelles

The SDS concentration is greater than the critical micelle concentration (cmc), so SDS is present as micelles. Micellar SDS bound to the tracking dye unstacks at high gel concentrations in disc electrophoresis; so a tracking dye that marks the moving boundary front at low gel concentrations fails to do so at high concentrations (source)

KCl causes SDS to precipitate

If you samples contain KCl you should dilute them or methanol precipitate them and resuspend them in 1X sample buffer. With low concentrations of KCl (<200 mM) you can run them on the gel but you should loaed every lane with sample buffer containing the same concentration of KCl (even if they are blanks). This will help the gel run a little less anomalously. (source)